New insights into centrosome imaging in Drosophila and mouse neuroepithelial tissues

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New insights into centrosome imaging in Drosophila and mouse neuroepithelial tissues Maria A. Rujano*, x, Renata Basto*, 1, Ve´ronique Marthiens*, 1 *Institut Curie, CNRS UMR144, Paris, France x Imagine Institute, Paris, France 1

Corresponding authors: E-mail: [email protected]; [email protected]

CHAPTER OUTLINE Introduction ................................................................................................................ 2 1. Imaging Centrosomes in Drosophila Neuroepithelial Cells......................................... 4 1.1 Materials ................................................................................................ 5 1.1.1 Reagents.............................................................................................. 5 1.1.2 Equipment ........................................................................................... 5 1.2 Methods ................................................................................................. 7 1.2.1 Larval staging ....................................................................................... 7 1.2.2 Dissection ............................................................................................ 7 1.2.3 Immunolabeling of fixed brains ............................................................. 8 1.2.4 Live imaging ........................................................................................ 9 2. High Spatial Resolution Imaging of Centrosomes in Neural Stem Cells of the Mouse Neocortex ................................................................................................. 10 2.1 Material ............................................................................................... 10 2.1.1 Reagents............................................................................................ 10 2.1.2 Equipment ......................................................................................... 12 2.2 Methods ............................................................................................... 12 2.2.1 Preparation of dorsal telencephalon explants....................................... 12 2.2.2 Immunolabeling of whole-mount cortical explants................................ 13 2.2.3 Imaging of centrosomes and mitotic spindles from an en face perspective ........................................................................................ 15 Conclusion ............................................................................................................... 16 Acknowledgments ..................................................................................................... 16 References ............................................................................................................... 17

Methods in Cell Biology, Volume 129, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2015.04.005 © 2015 Elsevier Inc. All rights reserved.

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Abstract The centrosome is the main microtubule-organizing center in animal cells. It participates in the assembly of a bipolar spindle that ensures accurate segregation of chromosomes during mitosis. Recently, mutations in centrosome genes have been identified in patients affected by neurodevelopmental disorders. In fact, the etiology of several neurodevelopmental pathologies seems to be linked to defects in the assembly of the mitotic spindle in the neural stem cell compartment during neurogenesis. Therefore, getting better insights into the structure and function/dysfunction of the mitotic spindle apparatus in an intact tissue environment is of utmost importance. However, imaging nanometer-scale structures like centrosomes and microtubule bundles within the depth of a tissue is still challenging. Here we describe two procedures to acquire high-resolution images on fixed tissues and to perform live imaging of microtubule-based structures in the neuroepithelia of the Drosophila brain and of the mouse neocortex. We take advantage of the accumulation of centrosomes and mitotic figures at the apical surface of these polarized tissues to improve the quality of staining and imaging. Both Drosophila and mouse models with centrosome dysfunction showed abnormalities in the neuroepithelium reminiscent of the ones described in brains of human patients. These observations have highlighted their value as model organisms to study the etiology of human neurodevelopmental pathologies.

INTRODUCTION Centrioles are cylindrical structures of 200 nm diameter and 500 nm length formed by triplets of microtubules arranged in ninefold symmetry (Bornens, 2002; Firat-Karalar & Stearns, 2014). Centrioles are embedded in and organize the pericentriolar material (PCM), the site of microtubule nucleation and anchoring (Fu & Glover, 2012; Lawo, Hasegan, Gupta, & Pelletier, 2012; Mennella, Agard, Huang, & Pelletier, 2014). A key step to investigate the function and dysfunction of centrosomes under normal and pathological conditions is to perform an in-depth characterization of their architecture and dynamics at different stages of the cell cycle (Chavali, Putz, & Gergely, 2014; Godinho & Pellman, 2014; Nigg & Raff, 2009). Nonetheless, imaging nanoscale centrosomal structures and the associated microtubule network in 3-dimensions (3D) within a tissue has remained challenging. The Drosophila neuroepithelium in the larval optic lobe is composed of symmetrically dividing neuroepithelial cells that generate asymmetrically dividing neural stem cells (NSCs) (Egger, Boone, Stevens, Brand, & Doe, 2007; Yasugi, Umetsu, Murakami, Sato, & Tabata, 2008) (Figure 1(A)). These cells form a monolayered pseudostratified columnar epithelial tissue (Rujano, Sanchez-Pulido, Pennetier, le Dez, & Basto, 2013) that displays typical apicobasal polarity within the larval optic lobe, and gives rise to all the neurons and glia of the adult optic lobe (Meinertzhagen & Hanson, 1993). The possibility of studying the mechanisms involved in cell fate transition from a proliferative (neuroepithelial) to a neurogenic (neuroblast) state, underscore the relevance of the optic lobe to study neurogenesis. Furthermore, the

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Introduction

FIGURE 1 Drosophila and mouse neuroepithelial tissue architecture. Drawings depicting the organization of embryonic neural stem cells within the neuroepithelium of Drosophila optic lobe (A) and mouse neocortex (B). Nuclei undergo interkinetic nuclear migration. This gives a pseudostratified appearance to the neuroepithelia. Centrosomes (red (gray in print versions) dots) and mitotic spindles always accumulate at the apical side of the tissue. (A) Drosophila neuroepithelial cells display a columnar shape and apicobasal polarity, with the apical side facing the outer part of the larval optic lobe and the basal side facing the inner part. Tissue width lies between 15 and 20 mm. (B) Mouse radial glial cells display an elongated bipolar morphology and apicobasal polarity, with processes keeping attachment to both basal lamina (outer) and ventricular lining (inner). Tissue width is increasing over neurogenesis, while the size of the ventricular lining is decreasing. En face view of the ventricular surface on the right-hand drawing illustrates the accumulation of mitotic spindles and interphase centrosomes from an apical point of view.

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Drosophila neuroepithelium is amenable to live imaging with high spatial and temporal resolution from both the apical and the lateral side and as such, it represents an ideal system to study centrosome architecture and dynamics in the context of a living developing tissue. The mouse embryonic neuroepithelium of the presumptive cerebral cortex is one of the best-documented model systems in mammals for studying the process of neurogenesis and related defects (Taverna, Gotz, & Huttner, 2014). Embryonic NSCs, also called radial glial cells, divide symmetrically to amplify the pool of progenitors and then asymmetrically to give rise to all neurons and glial cells populating functional adult brains (Temple, 2001). NSCs display an elongated shape spanning the whole thickness of the tissue. During the cell cycle, their nucleus moves up and down the depth of the tissue while their processes keep attachment to both basal and apical sides (Figure 1(B)) (Noctor, Flint, Weissman, Dammerman, & Kriegstein, 2001). Centrosomes remain closely attached to the apical side by an unknown mechanism. The apical surface of the neuroepithelium is in contact with the cerebrospinal fluid circulating in the third ventricle and is therefore often referred to as the ventricular surface. During interphase, one of the two centrioles templates the formation of a primary cilium at the apical side (Dubreuil, Marzesco, Corbeil, Huttner, & WilschBrauninger, 2007). Before mitosis, the nucleus reaches the ventricular surface where centrosomes accumulate. During mitosis, the two apical centrosomes organize the two poles of the spindle apparatus that participates in the faithful segregation of chromosomes in the two daughter cells. We take advantage of the apical centrosome location in NSCs all along the cell cycle to visualize them with high resolution in mouse cerebral cortical explants using en face view imaging. In the first part, we provide protocols for imaging centrosomes and microtubulebased structures in NSCs of the Drosophila optic lobe during larval stages by live imaging and/or immunolabeling of fixed tissues. In the second part, we describe methods to analyze centrosomes and mitotic spindles with high spatial resolution in embryonic NSCs of the mouse dorsal telencephalon. In particular, we explain how to perform whole-mount cortical explant analysis from an en face perspective on fixed tissues.

1. IMAGING CENTROSOMES IN DROSOPHILA NEUROEPITHELIAL CELLS The protocol described thereafter works for immunolabeling or live imaging of brains in larval and pupal stages, but NSCs are actively dividing only during 3rd instar larvae. Furthermore, the protocol is also suitable for immunolabeling and live imaging of cellular structures other than centrosomes. First, we describe how to collect larvae at the desired stages and how to dissect the tissue. Then, we explain how to fix and label centrosomes with antibodies and finally, we describe how to mount the tissue for live imaging (Figure 2).

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1. Imaging centrosomes in Drosophila neuroepithelial cells

1.1 MATERIALS 1.1.1 Reagents • • • • • • • • • •

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Drosophila larvae at developmental stage of interest Phosphate-buffered saline (PBS) (1X, pH 7.4) PBST 0.1% (PBS containing 0.1% Triton X-100) PBST 0.3% (PBS containing 0.3% Triton X-100) Normal goat serum (NGS, Invitrogen, PCN5000) Blocking solution (5% NGS v/v in PBST 0.3%) Paraformaldehyde 16% (Electron Microscopy Sciences, 15710) Paraformaldehyde 4% (diluted from paraformaldehyde 16% in PBST 0.1%) Primary antibodies diluted in PBST 0.3% One or more appropriate secondary antibodies diluted in PBST 0.3%. We recommend Alexa FluorÒ 488, 546, and 647 (Molecular Probes-Life technologies) Hoechst 33342 diluted at 0.5 mg/mL in PBST 0.3% (Hoechst 33342 trihydrochloride trihydrate, Molecular Probes-Life technologies, H1399; stock solution prepared at 10 mg/mL in DMSO and stored at 4  C) Mounting medium (1.25% n-propyl gallate (Sigma, P3130), 75% glycerol (bidistilled, 99.5%, VWR, 24388-295), 25% H2O). Prepared by mixing ingredients overnight (O/N) at 4  C and stored at 20  C Schneider’s Drosophila medium (Gibco, 21720-024) Heat-inactivated fetal bovine serum (FBS) (Gibco, 10500) Penicillin/streptomycin (Gibco, 15140) Live-imaging medium (Schneider’s Drosophila medium supplemented with 10% FBS, 100 units/mL Penicillin and 100 mg/mL streptomycin) Oil 10S Voltalef (VWR BDH Prolabo, 24627-188)

1.1.2 Equipment • • • • • • • • • • • • • •

Vials with cornmeal medium Dissecting microscope Silicon pad for dissection (plastic dish filled with SylgardÒ 184 silicone elastomer kit, Down Corning) Glass dishes (Nine-well Pyrex plate, Electron Microscopy Sciences, 71563) Forceps (two pairs; Dumont #5; Fine Science Tools, 11200-14) Micropipettes and tips Minutien pins (0.1-mm diameter, FST, 26002-10) Microdissecting pin holder (FST, 26018-17) Microscope slides Round cover glasses (12 mm ø, N 1, Marienfeld Lab Glassware, 01-115-20) Nail polish Orbital shaker Paintbrush Slide holder

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FIGURE 2 Imaging of Drosophila neuroepithelial cells. (A) Schematic representations of the successive steps of dissection required for isolating the Drosophila larval brain. (B) Larval brain mounting for imaging. The left-side panel shows a scheme for cleaning the brain before imaging. Right panel shows schemes of mounted samples for confocal imaging of fixed brains (top) and live imaging of fresh samples (bottom). (C) Example of confocal acquisition for centrosomes, microtubules, and actin staining.

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1. Imaging centrosomes in Drosophila neuroepithelial cells

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Glass bottom 35 mm dish (P35G-1.5-14-C, MatTek Corporation) Permeable membrane (Standard membrane kit, YSI)

1.2 METHODS 1.2.1 Larval staging As the developmental period in Drosophila is temperature-dependent, to facilitate reproducible staging of larvae, we recommend raising fly stocks and performing the staging at 25  C. In order to analyze centrosomes and cytoskeletal structures at different developmental stages in neuroepithelial cells, larvae are staged and the tissue is collected and analyzed at early, mid-, and late-third instar larvae (L3). 1. Place at least 20 adult flies (males and females) in vials with cornmeal medium at 25  C. 2. Collect eggs by flipping the flies from the vial after 2e4 h and keep the vial with eggs at 25  C. 3. Collect as many times as necessary depending on the fertility and viability of parental lines. 4. Select larvae for dissection of brains 72, 96, or 120 h after collection of eggs for early, mid-, and late-L3 larval stages, respectively. Optional: Collect only freshly hatched larvae in a 2e4 h time window and stage on cornmeal medium to early, mid-, and late-L3 larval stages (48, 72, or 96 h after larval hatching).

1.2.2 Dissection This is the general procedure for dissecting larval brains (Figure 2(A)). Critical: If dissection is followed by immunolabeling of fixed tissues, the dissection is performed in PBS. Alternatively, when preparing brains for live imaging, the dissection should be performed in the medium for live imaging.

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Centrosomes are depicted in red (light gray in print versions) and immunostained with Drosophila pericentrin-like protein (Plp), microtubules are in blue (gray in print versions) and immunostained with a-tubulin, actin is in red (light gray in print versions) and stained with phalloidin, and DNA is in gray labeled with Hoechst. (D) Example of spinning disc confocal live imaging of centrosomes and microtubules. Centrosomes are visualized with SPD::RFP (top panel) or Sas4::GFP (bottom panel) and microtubules are visualized with a-tubulin::GFP (top panel) or a-tubulin::RFP (bottom panel). Time is given in minutes. The neuroepithelium is imaged from the lateral side (top panel) and from the apical side (bottom panel). Fourdimensional z-stacks of 5e10 mm at 0.3 mm intervals were acquired every 30 or 60 s using an X60, NA 1.4 oil immersion objective. Scale bars: 5 mm

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1. Collect the larvae at the stage of interest from the medium or the walls of the vials with a PBS-soaked paintbrush or forceps and place them in one of the wells in a glass dish filled with PBS. Remove the fly food from the surface of the larvae carefully with a paintbrush. Subsequently, transfer the larvae into another well filled with fresh PBS to complete cleaning. 2. Transfer the larvae individually into drops of cold PBS or live-imaging medium placed on a silicon pad for dissection. 3. Tear larval body in the middle using the forceps and discard the posterior half. 4. Expose the brain, by holding the anterior half of the larvae by the mouth hook with one pair of forceps while turning the anterior inside out with the other pair of forceps. 5. Once the brain is located and while still holding the mouth hook, gently remove all other tissues from around the brain leaving only some imaginal discs attached. These ones are used to manipulate the sample without touching/damaging the brain. 6. Gently detach the brain from the mouth hook with the forceps and transfer to a clean drop of PBS or live-imaging medium by grabbing the tissue from one of the imaginal discs. A typical immunolabeling experiment requires 10e20 brains. Critical: To avoid tissue degradation, the time between dissection and fixation should not exceed 15 min. If necessary, dissect and fix brains in groups of 5e10. For live imaging, 1e4 brains can be mounted immediately after dissection.

1.2.3 Immunolabeling of fixed brains 1.2.3.1 Fixation 1. Using a P-20 micropipette, transfer the brains from the PBS drop, into a well of a glass dish that contains 200e500 mL of freshly prepared 4% paraformaldehyde. Tip: to avoid the tissue sticking to the microtip, before pipetting the tissue, pipet up and down on a solution of PBST or NGS. 2. Incubate the brains in the fixative on an orbital shaker with gentle agitation for 20 min at room temperature (RT, 20  C). Critical: the tissue should be completely submerged in fixative. If necessary, remove air bubbles with the help of forceps and/or fine mounted pins.

1.2.3.2 Staining 1. After fixation, wash the tissue in PBST 0.3% for 10 min with gentle agitation. Repeat this step 2 more times. Optional: Some antibodies may require a blocking step to minimize background. To block, add 100e500 mL blocking solution and incubate the fixed brains on an orbital shaker with gentle agitation for 30 min at RT. 2. Replace the PBST 0.3% with 50e200 mL primary antibody solution and incubate O/N at 4  C. Tip: to avoid evaporation, place the dish containing the brains on top of a water soaked Kimwipe inside a plastic box.

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1. Imaging centrosomes in Drosophila neuroepithelial cells

3. Remove antibody solution and wash in PBST 0.3% for 10 min with gentle agitation at RT. Repeat the washing step 2 more times. 4. Replace the PBST 0.3% with 50e200 mL secondary antibody solution and incubate O/N at 4  C. Optional: The incubation time in this step can be reduced to 3 h at RT but the antibody penetration greatly increases when performed O/N. 5. Remove antibody solution and wash in PBST 0.3% for 10 min with gentle agitation at RT. Repeat the washing step 2 more times. 6. Replace the PBST 0.3% with Hoescht solution and incubate for 3 h at RT with gentle agitation. 7. Remove the nuclear staining solution and wash with PBS for 10 min with gentle agitation at RT. Repeat the washing step 2 more times.

1.2.3.3 Mounting (Figure 2(B) and (C)) 1. Gently transfer one brain to an w7 mL drop of mounting medium on a microscope slide grabbing the tissue by one of the imaginal discs using the forceps. 2. Use fine mounted pins to remove the remaining imaginal discs and any other tissue attached to the brain (If not too big and is placed well separated from the brain, this excess tissue can be left on the slide). 3. Place the brain with the dorsal part facing the slide and ventral up. Use the fine mounted pins for this step. 4. Gently cover the brain with a round coverslip. 5. Soak excess mounting medium from the borders of the coverslip using a Kimwipe. 6. Seal the edges carefully with nail polish and place the samples on a dark slide holder. Optional: Up to three brains can be mounted on one slide. 7. Proceed to imaging. Perform imaging using a confocal microscope. For imaging of centrosomes on L3 larval brain cells, we recommend a 63 or 100 oil objective and a z-step of 0.2 mm Critical: Imaging should be done as soon as possible to get the best signal. After imaging, return the samples to a dark slide holder and keep them at 4  C.

1.2.4 Live imaging (Figure 2(B) and (D)) 1.2.4.1 Mounting 1. Use fine-mounted pins to remove imaginal discs and any other tissue attached to the brain. 2. Using a P-20 micropipette, gently transfer 1e4 freshly dissected brains to a 10 mL drop of medium on the center of a glass bottom 35 mm dish. Avoid transferring more than 5 mL of medium together with the brains. Tip: to avoid the tissue sticking to the microtip, before pipetting the tissue, pipet up and down live-imaging medium. 3. To image the neuroepithelium laterally, place the brain with the dorsal part facing the glass bottom. If imaging is to be done from the apical side, place the

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brain with the ventral part facing the glass bottom. Use the fine mounted pins for this step. 4. Cover the brains with a piece of permeable membrane small enough to fit on the space of the round glass bottom. 5. Seal the membrane edges with 10S Voltalef oil. 6. Proceed to imaging. Perform imaging using a spinning disc confocal microscope. For imaging of centrosomes in L3 larval brain cells, we recommend a 63 or 100 oil objective and a z-step of 0.2 mm. For centrosome dynamics studies, acquire images at intervals of 5e30 s maximum.

2. HIGH SPATIAL RESOLUTION IMAGING OF CENTROSOMES IN NEURAL STEM CELLS OF THE MOUSE NEOCORTEX First we provide a detailed protocol on how to dissect and fix the embryonic cortical wall of the presumptive cerebral cortex (Figure 3). Then, we provide detailed explanations on how to perform immunofluorescence staining of whole-mount cortical explants and how to image en face views of mitotic figures within the apical progenitor compartment.

2.1 MATERIAL 2.1.1 Reagents •

Mouse embryos collected between embryonic day 11.5 and 16.5 (referred to as E11.5 and E16.5, respectively) Nota Bene: All the solutions are prepared fresh the day of the experiment.

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PBS (1, pH 7.4) Paraformaldehyde 4%, prepared from a stock of PFA 16% (Electron Microscopy Sciences, 15710), diluted in PBS 1 PBST 1%, PBS 1 containing 1% Triton X-100 v/v (Sigma, T9284) and sodium azide 0.02% PBST 0.3%: PBS 1, 0.3% Triton X-100 v/v, sodium azide 0.02% Bovine Serum Albumin (BSA, FractionV-pH7, Euromedex, 04-100-811C) Blocking solution: 3% BSA w/v diluted in PBST 0.3% Primary and secondary antibodies are diluted in the blocking solution. We recommend the highly cross-adsorbed Alexa FluorÒ 488, Alexa FluorÒ 568, and Alexa FluorÒ 647 raised in goat for confocal imaging (Molecular Probes-Life technologies) DNA is stained with Dapi (40 ,60 -diamidino-2-phenylindole, Thermo Scientific, 46190). Dapi is diluted in the solution of secondary antibodies at a 3 mg/mL final concentration

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FIGURE 3 En face views imaging of centrosomes in the mouse neocortex. (A) Schematic representations of the successive steps of dissection required for isolating dorsal telencephalon explants from mouse embryonic brains before fixation and immunolabeling procedures. (B) Drawings explaining the mounting procedure to obtain apical side up dorsal telencephalon explants to image centrosomes in interphase and mitotic figures from an en face point of view. (C) Examples of en face view confocal acquisitions showing centrosomes (g-tubulin in red (light gray in print versions)), microtubules (a-tubulin in green (gray in print versions)), and DNA in blue (dark gray in print versions). The same region was imaged at different z-positions along the apicobasal axis. At the uttermost apical position (z ¼ 0 mm) (left-hand panel), the interphase centrosomes within the apical processes can be visualized (dashed white circle). In addition, the cleavage furrows are also visualized (white arrow). Just 4 mm above (z ¼ 4 mm) (right-hand panel) mitotic cells are visualized in different stages (white dashed circle surrounds one metaphase and red (light gray in print versions) dashed circle surrounds one anaphase). In the left-hand panels, note that centrosomes in interphase belong to cells at different stages of the cell cycle, i.e., one or two centrosomes can be visualized per apical process. In the right-hand panels, note that the parallel orientation of the mitotic spindle in relation to the ventricular lining enables the visualization of the two mitotic centrosomes at the same z-position. Scale bars: 8 mm.

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Mounting medium (1.25% n-propyl gallate (Sigma, P3130), 75% glycerol (bidistilled, 99.5%, VWR, 24388-295), 25% H2O). Prepared by mixing ingredients overnight (O/N) at 4  C and stored at 20  C Optional: For tissue pre-permeabilization before fixation, prepare a fresh solution of PBST 0.1%: PBS 1 containing 0.1% Triton X-100 v/v, respectively, and sodium azide 0.02%

2.1.2 Equipment • • • •

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Dissecting microscope One pair of scissors 8.5 cm straight (Fine Science Tools, FST, 14084-08), to collect uterine horns containing embryos by caesarean Two pairs of forceps (FST, Dumont #5, 11200-14) One pair of forceps angled up 7 mm (FST, Dumont #7, Dumoxel 11271-30), to hold embryos before chopping their heads off and pinch out brains and cerebral hemispheres One pair of fine scissors angled up, cutting edge 10 mm (FST, 15017-10), to chop the heads off embryos Silicon pad for dissection (plastic dish filled with SylgardÒ 184 silicone elastomer kit, Down Corning) One pair of fine scissors straight, cutting edge 3 mm (FST, 15000-00), to cut the skin and the skull of the heads and microdissect the median and ventral parts of the telencephalon Spatula type Chattaway (L:100  l:4 mm, Dutscher, 442180) or plastic dispensable micropipette (VWR, 612-1747), to handle and/or transfer dorsal telencephalon explants Microknife (Corneal/Scleral V-Lance Knives, Alcon Ophthalmic, 20G, 1.3 mm, 8065912001) to slice cortical explants 24-well plates for cell culture (TPP, 92024) Orbital shaker Micropipettes and tips Microscope slides Square cover glasses (18 mm diameter, No.1.5H, Marienfeld, 0117580) Nail polish

2.2 METHODS 2.2.1 Preparation of dorsal telencephalon explants 2.2.1.1 Collection of cerebral hemispheres 1. The observation of a vaginal plug is considered as day 0.5 of mating. Collect cerebral hemispheres when neurogenesis occurs between embryonic day 11.5 (E11.5) and embryonic day 16.5 (E16.5) for analysis. 2. Pregnant females are anaesthetized with isoflurane before being sacrificed by cervical dislocation. Collect embryos by caesarean with the help of 8.5 cm

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2. High spatial resolution imaging of centrosomes in neural stem cells

straight scissors. Remove them from the uterine horns and yolk sacs with a pair of forceps (#5). Hold the embryos with 7 mm angled-up forceps and chop heads off embryos with angled up fine scissors. Place the heads in a drop of PBS on a silicon plate for further dissection at RT (20  C). Note: Dissections are always performed at RT in order to prevent microtubule depolymerization before fixation.

2.2.1.2 Dissection of dorsal telencephalon explants (Figure 3(A)) The dorsal part of the telencephalon is isolated from other brain regions using the following dissection steps. 1. Use a pair of fine straight scissors to make a horizontal incision of the skin at the base of the forebrain. 2. Use the same scissors to cut the skin and the forming skull at the midline along the rostrocaudal axis. Perform the incisions superficially to prevent any damage of the underlying cerebral hemispheres. Care should be taken while dissecting E15.5eE16.5 brains since the skull is getting harder at these stages. Push away the skin and the skull laterally with a pair of forceps (#5) to uncover the cerebral hemispheres. 3. Pinch out each cerebral hemisphere from the rest of the brain with the #5 tweezers and orient the neocortical explant with the ventricular surface up in a drop of clean PBS. 4. Use tweezers (#5) to gently hold the cerebral hemisphere while cutting away the olfactory bulb, the ventral and median parts of the telencephalon with the fine straight scissors. Critical: Expose the ventricle surface of the dorsal telencephalon as much as possible at this step to enable a better access of the antibodies during immunolabeling. In that way, we can observe homogeneous staining of the whole surface area and a high reproducibility of our immunolabeling results. Notes: a. Perform the dissection steps as quickly as possible for a better preservation of the tissues. The whole dissection procedure should not exceed 45 min. b. The size of the ventricular surface is expected to decrease during late stages of neurogenesis as a consequence of depletion of the NSC pool. c. If the regional variability along the rostrocaudal and mediolateral axes is not an issue for the analysis, each cortical explant can be further sliced in four pieces maximum with the help of a microknife (V-Lance knife) on the silicon pad.

2.2.2 Immunolabeling of whole-mount cortical explants 2.2.2.1 General recommendations 1. Prepare all fixative solutions and buffers freshly the day of the experiment. 2. Perform the following steps by incubating each free-floating dorsal telencephalon explant in a well filled with the appropriate working solution. The use of

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24-well plates is recommended for the whole procedure, but 96-well plates can be used to reduce the amount of antibodies. 3. Fill and remove solutions from the wells with the help of a 1 mL blue pipet tip, or alternatively transfer explants from one well to another one with the help of a microspatula or a dispensable plastic micropipette. 4. Perform the whole procedure at RT. Incubate explants under gentle rotation (without the pre-extraction and fixation steps).

2.2.2.2 Fixation of dorsal telencephalon explants 1. Optional: Pre-extraction of the cytoplasmic pool of proteins before fixation improves the intensity of staining on centrosomes and other microtubule-based structures. For example, the organization of microtubule bundles within the spindle apparatus appears more clearly after pre-extraction of the pool of soluble a-tubulin. To perform this step, incubate the explants in a solution of PBST 0.1% for 2 min and wash them quickly 2 times in PBS. 2. Fix tissues in 20  C-precooled methanol for 10 min or alternatively in PFA 4% for 1 h at RT, depending on the primary antibody to be used. Wash the explants 3 times in PBS for at least 10 min before proceeding with immunolabeling. Note: It is highly recommended to perform the immunostainings immediately after methanol fixation. PFA-fixed explants can be stored at 4  C for one month in PBS 1 containing 0.02% azide before immunolabeling.

2.2.2.3 Immunofluorescent staining and mounting (Figure 3(B)) 1. Incubate fixed explants with 1 mL of PBST 1% for 15 min. 2. Perform the blocking step by incubating the explants in 1 mL blocking solution (3% BSA w/v diluted in PBST 0.3%) for 1 h at RT under gentle rotation. Use the same blocking solution to proceed for staining with primary and secondary antibodies. 3. Incubate the explants with primary antibodies (250 mL per well for 24-well plates, 50 mL per well for 96-well plates) O/N at RT under gentle agitation. Wash them 3 times with 1 mL blocking solution for at least 1 h. 4. Incubate the explants with secondary antibodies along with Dapi (staining of the DNA) O/N at RT, protected from light. Wash them 3 times with 1 mL blocking solution for at least 1 h and perform a last wash in PBS before mounting. 5. Place the explant in a drop of PBS on a glass slide for ease of manipulation and further orientation under a binocular microscope (Figure 3(B)). To obtain an en face view of the ventricular side of a cerebral cortex, place the explant as flat as possible with the ventricular surface up. Remove the excess of PBS with a 100 mL pipet yellow tip and add a drop of mounting

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medium onto the explant. Place a coverslip on the sample and seal it with nail polish. 6. Place the mounted samples at 4  C in a dark slide holder before imaging. Wait at least 24 h before imaging. The samples can be imaged until 1 month after immunolabeling.

2.2.3 Imaging of centrosomes and mitotic spindles from an en face perspective (Figure 3(C)) 2.2.3.1 Confocal imaging of centrosomes and mitotic figures 1. Image the ventricular side of dorsal telencephalon explants with a laser scanning confocal microscope. We can image centrosomes and mitotic spindles with the highest resolution using the Leica SP8, equipped with hybrid detectors (PICTIBISA Imaging Platform, Institut Curie) (Figure 3(C)). A 63 oil immersion lens (Leica  HCX PL APO 1.4 oil CS2) is used for all the acquisitions. If mitotic figures can be analyzed without any further zooming, we can get the best resolution for analyzing centrosomes and microtubule bundles within the mitotic spindle apparatus with an electronic zoom of 2, at a 1024  1024 pixels image resolution. 2. Interphase and mitotic centrosomes of NSCs remain at the ventricular surface and can be imaged at the same time using three-dimensional z-stacks of 8e10 mm at 0.2 mm intervals starting from the ventricular lining. Indeed, mitotic cells display a diameter between 8 and 10 mm depending on the mitotic stage and the embryonic age of the tissue. The ventricular lining can be easily identified by the accumulation of centrosomes at the apical surface. Centrioles are around 200 nm in diameter. Therefore, images taken using steps of 0.2 mm along the z-axis provide the best resolution of centrosomes (Figure 3(C)). Note: Regional variability can be observed along the rostrocaudal and mediolateral axes within the same cortical explant. For example, we have already noticed variability in the mitotic indexes (number of NSC in mitosis over total number of NSC in contact with the ventricle through their apical process). To check for and/ or get rid of such variability in our analysis, we perform each combination of staining using one whole cortical explant and image different area within the same explant (8e10 images acquired per explant).

2.2.3.2 Choice of antibodies for centrosomes and mitotic spindles staining 1. Centrosomes appear as small dots after fluorescent immunolabeling using antibodies, against centriole or PCM proteins (Figure 3(C)). In addition, mammalian cells display some satellite structuresdconsidered as reserve pools of centriole or PCMdwhich are spread around the centrosome and can be easily misidentified as the centrosome. Therefore, centrosomes should always be

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Imaging centrosomes in tissues

visualized by a combination of at least two centrosome markers. We recommend the following primary antibodies: a. g-tubulin antibody at 1/500 dilution (mouse monoclonal, clone GTU88, Sigma, T5326). This antibody stains the centriole wall and the PCM of centrosomes in methanol-fixed or PFA-fixed explants. b. pericentrin antibody at 1/500 dilution (rabbit polyclonal, Abcam, ab4448). This antibody stains the PCM of centrosomes in methanol-fixed or PFA-fixed explants. c. cdk5rap2 antibody at 1/500 dilution (rabbit polyclonal, Bethyl Laboratories, IHC-00063). This antibody stains the PCM of centrosomes in PFA-fixed explants. Alternatively, since centrosomes are microtubule-organizing centers during both interphase and mitosis, centrosome markers can be used in combination with a-tubulin. 2. To visualize the mitotic spindle apparatus, we recommend the following primary antibody: a. a-tubulin antibody at 1/1000 dilution (mouse monoclonal, clone DM1a, Sigma, T6199). This antibody stains microtubule bundles within the mitotic spindle apparatus, and especially microtubules nucleated at the centrosome, as well as the microtubule network below the plasma membrane.

CONCLUSION Here we described different approaches to study centrosomes within native tissue environments. These procedures were initially settled in our laboratory to analyze centrosome/spindle pole integrity and/or behavior in the NSCs of Drosophila and mouse mutant models for microcephaly-associated disorders (Marthiens et al., 2013; Rujano et al., 2013). They can now be used to get further insights into centrosome/spindle pole characteristics in relation to other neurodevelopmental disorders related to NSC dysfunctions.

ACKNOWLEDGMENTS Image acquisitions were performed on workstations of the PICT-IBiSA Lhomond Imaging facility of Institut Curie. The authors wish to thank V.Fraisier, L.Sengmanivong, and F.Waharte from the PICT-IBiSA Lhomond Imaging facility of Institut Curie for technical assistance. Work in our lab is supported by a grant from AICR (13e0170) and an ERC starting grant (Centrostemcancer 242598), as well as by the Institut Curie, the CNRS and INSERM. Our laboratory is a member of the CelTisPhyBio labex.

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References

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